Hanging Cell Culture Inserts

Millicell® Hanging Cell Culture Inserts are sterile, general purpose devices for the growth and differentiation of various cell types. Uniquely designed flanges suspend the insert in an off-center position within the culture plate well, facilitating pipetting by creating a larger space to one side. Millicell® hanging inserts make it possible to study both sides of the cell monolayer and are excellent tools in co-culturing and permeability assays.

Key Millicell® Hanging Cell Culture Key Highlights

For co-culturing and permeability assays

Unique design allows easier basolateral access than other hanging inserts with less risk of contamination

Available in 5 pore sizes and 3 diameters (for 24-, 12-, or 6-well plates) , including a 1 µm pore size that is optically transparent for better visualization by microscopy. The inserts are easily prepared for SEM and TEM visualizing techniques, and they are compatible with cellular and/or fluorescent stains.

Ordering Information

Characteristics

HA

CM

PCF

PET

Microscopically Transparent

No

Yes

No

1 µm only

Tissue Culture Treated

No

No

Yes

Yes

Membrane Thickness

120 µm

50 µm

10 µm

10 µm

Matrix/ECM Coatable

Yes

Yes

Yes

Yes

Membrane Types Available:

Biopore Membrane (Hydrophilic PTFE)For Low Protein-Binding, Live Cell Viewing, and Immunofluorescent Applications: Compared to other membrane matrices, this membrane exhibits little or no background when using fluorescent stains. It is also transparent to provide for optimum viewing of live cells. Biopore membrane is low protein-binding and requires extracellular matrix (ECM) coating for attachment dependent cells. ECMs are in the Proteins section of this handbook.

MF-Millipore Membrane (Mixed Cellulose Esters)For Exceptional anatomical and Functional Polarization and Growth of Attachment-dependent Cells without Matrix: The surfactant free, mixed cellulose esters membrane can be used for cell surface receptor, in vitro toxicology, microbial attachment and polarized uptake assays. When compared to plastic, cells had two- to three-fold higher densities and a more cuboidal morphology with rounded nuclei.

Isopore Membrane (Polycarbonate)For Growth of Attachment-dependent Cells without Matri: This track-etched, hydrophilic polycarbonate membrane is tissue culture treated to allow growth of attachment-dependent cells without the use of extracellular matrix (ECM). It is especially recommended for transport/permeability applications. The inserts are available in 5 pore sizes.

PET Membrane (Polyethylene Terephthalate)For Growth of Attachment-dependent Cells without Matrix: This track-etched, thin film membrane is translucent or microscopically transparent for better cell visualization and monitoring of the cell monolayer. It is tissue culture treated to promote cell attachment and growth.

Note: Depending on the cell lines and the nature of the co-culture, the researcher can decide which side of the membrane is seeded first. To maintain sterile incubations of cells seeded on the underside of the membranes, use sterile Petri dishes for the Millicell® single-well inserts and sterile feeder trays for Millicell® 24- and 96-well cell culture insert plates.

1. Using an optimized seeding density, seed the first cell type in either the apical wells or on the basolateral underside of the membranes. Refer to the recommended working volumes chart (see page 42) for appropriate apical volumes. For basolateral seeding volumes, use approximately 200 ìL for Millicell® 12 mm single-well inserts and Millicell®-24 insert plates, and approximately 30 µL for Millicell®-96 insert plates. 2. Incubate in a 37°C CO2 incubator for 1 to 4 hours to allow the cells to attach to the membrane. 3. Gently turn the Millicell® device over and seed the second cell type in either the apical well or on the basolateral underside of the membrane. 4. Incubate in a 37°C CO2 in the incubator for 1 to 4 hours to allow the second cell type to attach. 5. Add appropriate volume of media to the apical or basolateral chambers and return to incubator.

D. Day 4-8 1. Feed ESC on MEF feeder layer with fresh ESC media or pass cells, at a 1:2 ratio, if required. (After thawing ESC, 2–3 passages are preferred before seeding onto a Millicell®-24 or Millicell® 96-well plate. Both cell types are lifted at once and passed on to a new T-75 containing inactivated MEF.) ESC colonies grown on Millicell®-96 cell culture insert 1.0 PET membrane, stained for alkaline phosphatase activity, after culturing via indirect co-culture with ESC in apical well, at a 200 cell per well seeding density, and MEF in the single-well feeder tray.

G. Day 12 and Day 14 1. Feed ESC and MEF indirect co-culture with ESC media.

H. Day 16 1. Analyze alkaline phosphatase activity to demonstrate that ESC is undifferentiated with an alkaline phosphatase detection kit.

Note: This protocol is designed to grow undifferentiated embryonic stem cells in an indirect co-culture with the fibroblast feeder layer. Although it is targeted for use with Millicell®-24 and Millicell®-96 plates, this protocol can be used with Millicell® single-well inserts as well.

ECM Coating Protocols Coating of membranes and plastic surfaces with extra cellular matrices (ECMs) promotes cell attachment and monolayer formation. We have developed protocols for four types of ECMs on Millicell®-CM inserts. They are also useful for growing cells on plastic feeder trays in co-culture experiments.

Fixation and Staining ProtocolsMillicell® cell culture inserts, 24-well plates and 96-well plates are designed to support all fixation, staining and immunostaining procedures in a single device. Cells can be visualized by stereoscopic microscopy, phase contrast microscopy, or fluorescent methods. Many staining procedures employ a fixation step first. Fixation is required to stabilize sub-cellular morphology and prevent degradation of antigens during subsequent staining procedures. The following are examples of fixing and staining protocols.

If the chemicals are compatible with the membrane but not the polystyrene housing, remove the membrane from the housing before adding the chemical.

Unless otherwise stated, the chemicals listed are at maximum concentration. If the plastic housing and/or membranes are not compatible with the maximum concentration, they might be compatible at lower concentration.

Immunocytochemical staining is a technique employing fluorescently labeled antibody, by which cells can be localized, labeled, and examined via fluorescent microscopy.

Materials and Reagents

Millicell® Cell Culture Inserts and Insert Plates

Sterile Phosphate Buffer Saline (PBS)

Methanol, 100%

Glycerol

FITC-conjugated antibody

1% BSA in PBS

Glass slides

Method 1. Aspirate cell culture media and wash Millicell® inserts or plates gently on both sides with PBS. Incubate for 5 minutes and repeat 2–3X. Consult the recommended working volumes table for appropriate volumes. 2. Add fixative solution (e.g. methanol) for 5–10 minutes. It is not required to treat the underside of the membrane with fixative. Incubate according to protocol instructions. 3. After treatment, aspirate fixative and fill filter wells with washing buffer. Repeat 2–3X in order to fully remove the fixative solution from both sides of the filter membrane. Do not allow cells to dry. 4. Dilute primary antibody according to vendor recommendations. In order to obtain best results, it is recommended that optimal working dilutions be determined by the user. If permeabilization is required (such as for cytoplasmic or nuclear antigens), saponin can be added to the solution at a concentration of 0.1% 5. Add antibody solution to each well then incubate at recommended temperature (typically room temperature or 4°C) with mild shaking or rocking to assure that solution wets out entire filter surface. If antibody is fluorescently labeled (direct labeling), cover plate with foil to protect from light. 6. Aspirate antibody solution and wash both sides of membrane as indicated in Step 1 to remove all unbound antibody. 7. If performing indirect labeling with a secondary antibody, repeat steps 4 through 6 with the secondary antibody. For visualization using fluorescent antibodies, continue to Visualization and Microscopy procedures. For enzyme linked assays (HRP, etc.), follow vendor procedures for developing.

The Microscopic Examination of Samples Can Be Performed in Three Modes: 1. Viewing from below the plate (through transparent PET or CM membranes) Millicell® devices using PET or CM membrane have been designed to allow visualization of cells from below using an inverted microscope. For viewing live cells, microscopic observations can be made through the receiver or plastic plate containing media. In order to focus on the cells, the microscope objective (typically 5–20X) must have an appropriate working distance. (For objective specifications, visit the websites listed in the Microscope Objective Information section.) Fixed cells that do not require to be visualized in media can be viewed directly without the receiver plate. However, care should be taken not to contaminate the objective with liquid residue (media, mounting fluid) on the membrane. 2. Viewing from above the plate (Millicell® 6-well inserts, Millicell®-24 cell culture insert plates) Some cell culture platforms can allow the cells to be viewed in a conventional microscope directly from above using low magnification. Cells can be visualized through the lid to maintain sterility or with the lid removed for fixed cells or when maintenance of sterility is not required. Working distances of the objective must be longer when reading from above compared to when reading from below. If using immunofluorescence, it is recommended to use a mounting fluid that contains an anti-fade additive to prevent photobleaching. 3. Visualizing membranes on microscope slides (for higher magnification or withobjectives with short working distances [less than 2 mm]) The membrane can be removed from each well for microscopic evaluation. This allows for higher magnification examination and storage of the slides for future use.

For visualizing from above the membrane, typically 5–20X objectives are used that have at least a 13.59 mm (A) or a 18.03 mm (B) working distance when viewing without or with the lid, respectively. For visualizing from below the membrane, 5–20X objectives are used that have at least a 2 mm (C) working distance.

To prepare membranes on microscope slide 1. Remove the membrane from the well using a sharp scalpel to make a small incision in the edge of the membrane. Carefully cut along the inside of the well wall for approximately one quarter of the well diameter. Using forceps (Millipore cat. no. XX62 000 06), carefully hold the membrane while continuing to cut around the well diameter to remove membrane. Alternatively, a cork borer may be used to remove the membrane. Note: Use care to prevent membrane from curling. 2. Place the membrane disk, cells facing up, onto a microscope slide. 3. Add 50 µL mounting fluid to the membrane disk and allow it to wet out in order to prevent bubbles under the disk. 4. Slowly lower a cover slip onto the membrane at an angle to allow air bubbles to be removed.

Microscope Objective Information Information regarding microscope objective magnification power and working distances can be obtained from individual optical dealers or from the microscope vendors:

Ethanol Concentration Kit (%)

Time (minutes)

30

15

50

15

70

15

95

15

100

3 x 15

Note: Dehydration of Millicell®-HA units should be performed in a metal pan that will be used as the embedding tray due to the tendency of the cellulosic membranes to be less rigid during the dehydration process. Attempts to transfer the membranes during these steps could lead to mechanical damage to the cells.

6. For infiltration, EPON812, an EDPON substitution, or LX112 is suitable for both devices (do not use Spurr’s). The following is a general infiltration scheme:

Ethanol Concentration Kit (%)/Tray (% Plastic)

Time (minutes)

75/25

30 on a shaker

50/50

30 on a shaker

0/100

30 each/3x on a shaker

0/100

Overnight

Note: It is not necessary to use any other agent, such as propylene oxide, with plastic. Propylene oxide will dissolve the cellulosic filters. In addition, the standard inversion/rotation of specimens used in these steps is not advised since either (1) damage to the cell layer or (2) stretching of the cellulosic filter may occur. Mild shaking on a gel shaker apparatus is sufficient for successful infiltration.

Note: Before the next step the membrane must be detached from the surrounding plastic ring. Sometimes this will occur without manipulation since the EPON may loosen the membrane-to-ring bond. If this does not occur, use a sharp scalpel or a cork borer and cut the membrane. It may also help to cut the membrane over a 47 mm filter support disk. Under no circumstances should the membrane be left attached to the ring during polymerization.

7. Transfer to fresh plastic and polymerize at 68°C overnight.

B. Sectioning Notes 1. Nitrocellulose (HA), polycarbonate (PC) and polyethylene terepthalate (PET) membrane: These membranes can be sectioned in any plane without difficulty. 2. CM (Biopore) membrane: The Biopore membrane must be processed in one of two ways based on the final thickness of the section.